Significance
The generation of haploids is the most powerful means to accelerate the plant-breeding process. We elucidated whether point mutations in the centromere-specific histone H3 variant CENH3 could be harnessed for the induction of haploids. We identified plants with impaired centromere loading caused by a mutation in the centromere-targeting domain (CATD). The same mutation results in reduced loading of CENH3 in transgenic Arabidopsis and sugar beet. Arabidopsis plants carrying this single point mutation in wild-type CENH3 were used as haploid inducers. Because the identified mutation site is highly conserved and because point mutations can be generated by mutagenesis or genome editing, the described method offers opportunities for application in a wide range of crop species.
Keywords: haploid induction, CENH3 mutant, plant breeding, CENH3 loading, chromosome elimination
Abstract
The chromosomal position of the centromere-specific histone H3 variant CENH3 (also called “CENP-A”) is the assembly site for the kinetochore complex of active centromeres. Any error in transcription, translation, modification, or incorporation can affect the ability to assemble intact CENH3 chromatin and can cause centromere inactivation [Allshire RC, Karpen GH (2008) Nat Rev Genet 9 (12):923–937]. Here we show that a single-point amino acid exchange in the centromere-targeting domain of CENH3 leads to reduced centromere loading of CENH3 in barley, sugar beet, and Arabidopsis thaliana. Haploids were obtained after cenh3 L130F-complemented cenh3-null mutant plants were crossed with wild-type A. thaliana. In contrast, in a noncompeting situation (i.e., centromeres possessing only mutated or only wild-type CENH3), no uniparental chromosome elimination occurs during early embryogenesis. The high degree of evolutionary conservation of the identified mutation site offers promising opportunities for application in a wide range of crop species in which haploid technology is of interest.
The generation and use of haploids is one of the most powerful biotechnological means to accelerate the breeding process of cultivated plants. The advantage of haploid plants for breeders is that homozygosity can be achieved at all loci in a single generation via whole-genome duplication, without the need of selfing or backcrossing over many generations as conventionally required to obtain true-breeding lines. Haploids can be obtained in vitro or in vivo, although many species and genotypes are recalcitrant to these processes (reviewed in ref. 1). Alternatively, substantial changes to centromeric histone H3 (CENH3), such as replacing the hypervariable N-terminal tail of CENH3 with the tail of conventional histone H3 and fusing it to GFP (producing “tailswap-cenh3”), or complementing the cenh3.2-null mutant with homologs from the mustard family CENH3s creates haploid inducer lines in the model plant Arabidopsis thaliana (2–4). Haploidization occurred only when such a haploid inducer was crossed with a wild-type plant. The haploid inducer line proved to be stable upon selfing, suggesting that competition between modified and wild-type centromeres in the developing hybrid embryo results in the inactivation of the centromeres from the inducer parent. Consequently, chromosomes from the inducer parent are lost, and progeny can be recovered that retain only the haploid chromosome set of the wild-type parent.
Because CENH3 is almost universal in eukaryotes, this method has the potential to produce haploids in any plant species. To elucidate whether, in addition to the severe conformational change using the CENH3-tailswap (2, 3), nontransgenic-induced minimal mutations in endogenous CENH3 also could affect the centromere function for haploid induction, we screened a population of barley (Hordeum vulgare) produced by ethyl methanesulfonate-induced targeting of local lesions in genomes (TILLING) (5); this diploid species has two functional variants of CENH3, αCENH3 and βCENH3 (6, 7). Assuming that either CENH3 variant can compensate for the absence of the other, viable offspring should be observed in the presence of a loss-of-function allele in one of the two CENH3 variants. A total of 25 TILLING mutants were identified for both barley CENH3 genes (Table S1). The functionality of mutated CENH3s of homozygous M2 individuals of nine TILLING lines carrying nonsynonymous mutations was determined by immunostaining the centromeres with barley CENH3 variant-specific antibodies. αCENH3 and βCENH3 signals at centromeres were revealed in all but one mutant TILLING genotype 4528 (called “Hvßcenh3 L92F”) carrying a homozygous leucine-to-phenylalanine substitution at amino acid 92. It showed no centromeric βCENH3 signals in mitotic, meiotic, or interphase cells, and only minor βCENH3 signals were observed in the nucleoplasm outside the centromeres (Fig. 1). Because no obvious differences in the transcription levels of both CENH3 genes were found in wild-type CENH3 and Hvßcenh3 L92F (Fig. S1), the centromeric loading of the mutated βCENH3 seems to be impaired. Centromeres without βCENH3 are sufficient for mitotic centromere function, because no obvious chromosome segregation defects, such as anaphase bridges or changes in endopolyploidy, could be found (Fig. S2).
Table S1.
Characterization of barley αCENH3 and βCENH3 TILLING lines
| Gene | Plant ID | Mutation (position at gene, exon, or intron) | Type of mutation (position at protein) |
| αCENH3 | 1–41 | G→A (57-E1) | Silent K→K (19) |
| 1–39 | G→A (71-E1) | Missense S→N (24) | |
| 1–05 | C→T (342-E3) | Missense A→V (48) | |
| 10662–1 | G→A (345-E3) | Missense R→Q (49) | |
| 1–23 | A→T (358-E3) | Silent P→P (53) | |
| 15125–1 | G→A (371-E3) | Missense G→R (58) | |
| 11586–1 | G→A (762-E4) | Silent R→R (66) | |
| 10260–1 | G→A (793-E4) | Missense A→T (77) | |
| 12673–1 | G→A (813-E4) | Silent K→K (83) | |
| 13075–1 | C→T (841-E4) | Missense P→S (93) | |
| 15523–1 | A→T (1892-E6) | Silent A→A 1(38) | |
| 11917–1 | C→T (1907-E6) | Silent I→I (143) | |
| 15701–1 | G→A (1916-E6) | Silent K→K (146) | |
| βCENH3 | 12049 | C→T (171-E1) | Silent I→I (57) |
| 2485 | C→T (230-I1) | Silent noncoding | |
| 6899 | C→T (252-I1) | Silent noncoding | |
| 11787 | G→A (265-I1) | Silent noncoding | |
| 3589 | G→A (287-I1) | Silent noncoding | |
| 11643 | C→T (326-I1) | Silent noncoding | |
| 11872 | C→T (369-I1) | Silent noncoding | |
| 9779 | A→T (433-E2) | Missense K→M (79) | |
| 15244 | G→A (434-E2) | Silent K→K (79) | |
| 3583 | C→T (465-E2) | Missense L→F (90) | |
| 4528 | C→T (471-E2) | Missense L→F (92) | |
| 11301 | G→A (572-I2) | Silent noncoding |
Intron/exon model of barley CENH3s. Positions of only missense mutations are indicated by arrows. (A) αCENH3. (B) βCENH3.
Fig. 1.
Centromeres of barley TILLING line 4528 lost βCENH3. Chromosomes of wild-type and homozygous TILLING line 4528 after immunostaining with antibodies specific for αCENH3 (green) and βCENH3 (red). Note the absence of βCENH3-specific signals in line 4528.
Fig. S1.
Transcription level of αCENH3 and βCENH3 in wild-type (cv Barke) and TILLING line 4528 with mutated βCENH3. The relative expression level of αCENH3 and βCENH3 was measured in different tissues of barley using specific primers (Table S1). cDNA was prepared from total RNA, and gene expression levels were normalized to GAPDH. Each value represents the mean ± SD (n = 3). 7, 7 d after pollination; 14, 14 d after pollination.
Fig. S2.
The phenotypes of wild-type and homozygous TILLING line 4528 barley plants are similar. (A) Relative proportion of 2C, 4C, 8C, and 16C leaf nuclei determined by flow cytometry. (B) The cycle values indicating the mean numbers of endoreduplication cycles per nucleus (32) in wild-type 1–3 and in TILLING line 4528 lines 1–3 plants. (C) Growth habit of TILLING line 4528 (Left) and wild-type (Right) plants.
The potential of Hvßcenh3 L92F to act as a haploid inducer was tested. The analysis of 577 F1 plantlets obtained from crosses involving the Hvßcenh3 L92F mutant as the maternal (35 spikes) or paternal (22 spikes) partner for wild-type barley spikes did not reveal any haploid or otherwise hypoploid plants, whereas all 18 plantlets derived from nine wild-type barley spikes pollinated with Hordeum bulbosum (as a positive control for the procedure used to induce uniparental genome elimination) proved to be invariably haploid. This result indicates that the Hvßcenh3 L92F mutation in the presence of native αCENH3 is not sufficient for chromosome elimination during early zygotic embryogenesis.
The Hvßcenh3 L92F mutation is located in the CENH3 centromere-targeting domain (CATD), defined by loop 1 and the α2 helix of the histone fold domain (Fig. S3). This domain has been shown to be required for centromere loading of CENH3 by Scm3/HJURP chaperons in nonplant species (8, 9). The corresponding position in human CENH3, L91, is essential for CENH3 localization into the centromeres after binding with chaperones (8). CENH3 chaperons are highly variable among different organisms (10), and no analog for plants has been identified yet. However, the possibility that this domain in plants also mediates interaction with a chaperon cannot be excluded.
Fig. S3.
The CENH3 mutation is located in an evolutionarily highly conserved C-terminal CENP-A targeting domain (CATD) defined by loop 1 and helix 2 of the histone fold domain. Shown are multiple alignments of partial sequences of the CENH3 proteins from different monocot (Hordeum vulgare_βCENH3: JF419329; Hordeum vulgare_αCENH3: JF419328; Hordeum bolbusom_βCENH3: JF419330; Hordeum bolbusom_αCENH3: GU245882; Allium cepa: BAL45432; Avena sativa: AB981584.1; Oryza sativa: AY438639; Saccharum officinarum: CA127217; Sorghum bicolor: XM_002441245.1; Triticum aestivum: JF969286.1; and Zea mays: AF519807) and eudicot (Arabidopsis thaliana: AF465800; Beta vulgaris (this study); Brassica juncea: BAF49728; Brassica napus: ACZ04984.1; Brassica rapa: NM_001302028.1; Daucus carota: KJ201903; Glycine max: FK014964; Gossypium arboreum: KP177465; Gossypium hirsutum: KP177475.1; Gossypium raimondii: KP177464.1; Lepidium virginicum: BAF49732; Nicotiana tabacum: AB366153.1; Raphanus sativus: BAF49733.1; Solanum lycopersicum: XM_010328624-1; and Solanum tuberosum: XM_006339625.1) plant species and nonplant species (Drosophila melanogaster: AY126930; and Homo sapiens: AAP36900). The single point mutation in barley βCENH3 is indicated by the black arrow.
To determine whether the CATD mutation caused the observed impaired centromere loading, YFP was N-terminally fused to the coding sequence of A. thaliana CENH3 with an L→I or L→F exchange (L130I or L130F) at the corresponding position in A. thaliana CENH3. Double immunolabeling of transgenic A. thaliana with anti–wild-type CENH3 and anti-GFP revealed significantly reduced centromere targeting, especially of the L-to-F mutated CENH3s compared with wild-type CENH3 fused to YFP (Fig. 2A). In addition, the functional significance of the identified mutation was assayed in sugar beet (Beta vulgaris). RFP reporter constructs containing the cDNA of sugar beet CENH3 with an L106I or L106F exchange (corresponding to amino acid position 92 of barley; Fig. S3) were generated and used for stable transformation of sugar beet cells. Again, centromere targeting of the two mutated CENH3s was reduced compared with the endogenous CENH3 fused with RFP in callus and leaf nuclei of regenerated plants (Fig. 2B). In both species, the L-to-F exchange resulted in a stronger effect than the L-to-I exchange, likely because of improper function or folding of the protein as the result of steric challenges imposed by the additional aromatic group provided by phenylalanine (11). These results indicate impaired centromere targeting of CENH3 by a point mutation in the conserved amino acid sequence of monocot and eudicot species.
Fig. 2.
Characterization of the CENH3 point mutation in A. thaliana and B. vulgare. (A) Reduced centromere targeting of L130-mutated CENH3 in A. thaliana. (Upper) Quantification of centromere colocalization patterns (complete, partial, and no) in flower bud nuclei of wild-type A. thaliana (502 nuclei from three plants), cenh3 L130I (543 nuclei from four plants), and cenh3 L130F (1,105 nuclei from seven plants). (Lower) Representative double immunostaining patterns. Construct and endogenous wild-type CENH3 were detected with anti-GFP and anti-CENH3 antibodies, respectively. (B) Reduced centromere targeting of L106-mutated CENH3 in B. vulgaris. (Upper) Quantification of centromere colocalization patterns (complete, partial, and no) in callus and leaves of plants transformed with RFP reporter of wild-type CENH3 (230 nuclei from five plants) and Bvcenh3 L106I (160 nuclei from two plants) or Bvcenh3 L106F (350 nuclei from five plants) constructs. (Lower) Representative centromere patterns. (C and D, Upper) Distribution of CENH3 in Atcenh3 L130F-complemented cenh3.1-null mutants. Quantification of centromere colocalization patterns (strong, weak, or dispersed) in leaf nuclei of wild-type (1,071 nuclei from three plants), cenh3 L130F-1 (2,666 nuclei from six plants), cenh3 L130F-2 (1,157 nuclei from three plants), and cenh3 L130F-3 (1,656 nuclei from four plants) (C) and in sperm nuclei of wild-type (1,121 nuclei from three plants), cenh3 L130F-1 (1,918 nuclei from five plants), cenh3 L130F-2 (1,321 nuclei from three plants), and cenh3 L130F-3 (1,201 nuclei from three plants). (C and D, Lower) Representative anti-CENH3 distribution patterns. Double asterisks indicate 1% significance versus CENH3 wild type; single asterisks indicate 5% significance versus CENH3 wild type. (E) Flow histograms of diploid and haploid A. thaliana plants together with representative nuclei after FISH using an A. thaliana centromere-specific probe (in red).
To test for haploid inducer ability in A. thaliana, a genomic CENH3 construct previously used for functional complementation of cenh3.1 homozygous knockout plants (2) was L130F mutated and used to transform heterozygous cenh3.1 knockout plants (2). Lines with one, two, or three Atcenh3 L130F transgene insertions were selected (Fig. S4A). Consistent with the result obtained with the reporter cenh3 L/F construct in A. thaliana, and sugar beet, a high proportion of Atcenh3 L130F-complemented cenh3.1-null mutants displayed limited centromeric anti-CENH3 signals in diploid leaf and haploid sperm interphase nuclei (Fig. 2 C and D). The single insertion line Atcenh3 L130F-1 revealed the highest proportion of nuclei with impaired CENH3 distribution. Unlike the tailswap-CENH3 haploid inducer (2, 3), the phenotype, meiosis, and seed setting of homozygous Atcenh3 L130F lines were almost unaffected (Fig. S4 B and C). Thus, despite a diminished centromere loading of Atcenh3 L130F, mitosis and meiosis work sufficiently well to produce diploid offspring upon selfing. When Atcenh3 L130F-1 plants were pollinated with wild-type A. thaliana, 4.8% of total F1 seedlings were haploid (24.1% and 2.8% haploids of F1 seedlings grown from shriveled and normal seeds, respectively) and possessed only the chromosomes of the wild-type parent (Tables 1 and 2 and Figs. S5 and S6). In addition, plants derived from shriveled seeds were more often aneuploid or mixoploid. The same combination, including the other Atcenh3 L130F lines, resulted in diploids and aneuploids only (Tables 1 and 2). The reciprocal cross did not generate haploid plants.
Fig. S4.
Analysis of Atcenh3 L130F A. thaliana lines. (A) Determination of the Atcenh3 L130F transgene copy number by Southern blot. DNA blot [T2 plants of L130F-1 (two plants), L130F-2 (three plants), and L130F-3 (three plants)] were probed with a labeled hygromycin resistance gene. (B) Growth habit of wild-type and Atcenh3 L130F-1. Insets show the corresponding flower phenotype. (C) Analysis of meiosis, pollen quality, seed phenotype, germination frequency, and ploidy. The asterisk indicates the occurrence of lagging chromosomes or micronuclei.
Table 1.
Seed analysis of offspring derived from the reciprocal cross Atcenh3 L130F × wild-type A. thaliana
 |
Shading emphasizes the different crossing direction.
Table 2.
Ploidy analysis of offspring derived from the reciprocal cross Atcenh3 L130F × wild-type A. thaliana
 |
Seedlings were derived from seeds with a normal or shriveled phenotype. Shading emphasizes the different crossing direction.
Two hundred ten seedlings out of 307 germinated seeds were analyzed.
One hundred seedlings out of 129 germinated seeds were analyzed.
Fig. S5.
Haploid plants have only a wild-type CENH3 allele. Genotyping by dCAPS PCR. The Atcenh3-1 L130F mutant allele is not cut (215 bp); the CENH3 wild allele is cut (191 and 24 bp). cenh3-1/cenh3-1, complemented plant by genomic construct of cenh3 L130F; CENH3/cenh3-1, hybrid between Atcenh3 L130F and a wild-type plant. CENH3/CENH3, wild-type plant.
Fig. S6.
Scanning electron micrographs showing phenotypes of F1 seeds derived from the Atcenh3 L130F1 × wild-type A. thaliana cross.
To test whether the efficiency of the haploid inducer correlates with the total amount of CENH3, a comparative Western blot analysis was conducted. Indeed, lines L130F-1 and -2 possessed less total CENH3 than wild-type and L130F-3 plants (Fig. S7). The reduced amount of nuclear CENH3 in both lines could be caused either by efficient proteasome-mediated degradation that prevents promiscuous misincorporation of CENH3 throughout chromatin (12) or reduced nuclear transport of modified CENH3. Our results suggest that, depending on the amount of CENH3, a single point mutation in CENH3 is able to generate a haploid inducer in species carrying no more than one CENH3 variant.
Fig. S7.
Relative amount of CENH3 in different Atcenh3 L130F lines. The relative amount of CENH3 (CENH3/histone H3 signal intensity ratio) was determined by quantitative Western blot analysis with A. thaliana CENH3-specific antibodies and histone H3-specific antibodies (n = 3 plants of each line). Error bars indicate the 95% confidence interval. Double asterisks indicate 1% significance versus wild type.
We propose a possible model of how the process of uniparental chromosome elimination works in inducer cenh3 × wild-type CENH3 hybrid embryos (Fig. 3). It is likely that egg cells derived from haploid inducers contain less CENH3 than eggs derived from wild-type plants or contain a reduced amount of an unknown “CENH3-transgeneration required signature.” According to studies performed with a wild-type CENH3-GFP reporter, A. thaliana sperm nuclei but not egg cells are marked with CENH3 (13). However, it is possible that residual amounts of maternal CENH3 generating a “centromeric imprinting” are transmitted to the progeny. Within a few hours after fertilization paternal wild-type CENH3 also is actively removed from the zygote nucleus, and centromeric reloading of CENH3 in the zygote occurs at the 16-nuclei stage of endosperm development in A. thaliana (13). In embryos undergoing haploidization, centromeric reloading of the maternal chromosomes is impaired or delayed, causing chromosomes to lag at anaphase because of centromere inactivity. Subsequently, micronucleated haploid inducer chromosomes will degrade, and a haploid embryo will develop, as demonstrated for unstable interspecific hybrid embryos (14). In contrast, in a noncompeting situation in which centromeres possess only mutated or wild-type CENH3, the timing of centromere assembly is similar during early embryogenesis, and no chromosome elimination occurs.
Fig. 3.
Model explaining the uniparental chromosome elimination in haploid inducer CENH3 × wild-type CENH3 intraspecific hybrid embryos (Left) compared with embryos originating from A. thaliana wild-type crossings (Right). (A) Diploid homozygous haploid inducer and wild-type CENH3 parents produce haploid gametes during meiosis. (B) Egg cells derived from the haploid inducer likely contain either less CENH3 than eggs derived from wild type or a reduced amount of an unknown CENH3-transgeneration required signature. (C) After fertilization, the paternal wild-type CENH3 is actively removed from the zygote nucleus. (D) Centromeric reloading of wild-type CENH3 in the zygote occurs at the 16-nuclei stage of endosperm development in A. thaliana. (E and F) In embryos undergoing haploidization, centromeric reloading of the maternal chromosomes is impaired or delayed causing (i) lagging chromosomes and (ii) subsequent micronuclei formation because of centromere inactivity (E). Subsequently, micronucleated haploid-inducer chromosomes will degrade, and a haploid embryo will develop (F). Embryos contain paternal-derived chromosomes in the background of maternal-derived cytoplasm.
In summary, the CATD of the CENH3 gene was mutated in barley by a single-nucleotide exchange leading to impaired centromere loading of βCENH3. The effect was reproduced in transgenic A. thaliana and sugar beet exhibiting the same CENH3 mutation. Thus, genotypes carrying the described L130F mutation or a different amino acid switch at this position or at another positon within CATD could be developed into a general instrument for haploid induction in a wide range of eudicot and monocot species. Finally, because single amino acid mutations can be generated by chemical mutagenesis, the entire process of haploidization via application of a haploid inducer line is nontransgenic. Alternatively, haploidy inducers could be generated by genome editing (15) without any modification of the genetic background.
Materials and Methods
Hordeum vulgare.
Mutagenesis of barley αCENH3 and βCENH3 by TILLING.
We screened a TILLING population of 7,979 ethyl methanesulfonate-treated (EMS) diploid barley (H. vulgare) plants of cv. Barke (5) to identify mutant alleles of αCENH3 and βCENH3. Four and three primer combinations (Table S2) were used to amplify all exons and part of the corresponding introns of the αCENH3 and βCENH3 variants respectively, by using PCR with a heteroduplex step as described earlier (5). PCR products were digested with the dsDNA Cleavage Kit and analyzed using Mutation Discovery and Gel-dsDNA reagent kits on the AdvanCETM FS96 system according to the manufacturer’s guidelines (Advanced Analytical).
Table S2.
Primer sequences used in this study
| Primer name | Sequence (5′ to 3′) |
| Hv_aCENH3_EX1+2+3_F | AGGCAGGGTCTCAATTCCTT |
| Hv_aCENH3_EX1+2+3_R | GTCCCATCATCCATCGTCTT |
| Hv_aCENH3_EX4+5_F | CCCACTTCCTTGTTGTGGAC |
| Hv_aCENH3_EX4+5_R | GGCGATAAATGTATCTTGCATTC |
| Hv_aCENH3_EX6_F | TGGTAGCAACCAGAGCTACG |
| Hv_aCENH3_EX6_R | ACTGGCATGTTTCCTTCTGC |
| Hv_aCENH3_EX7_F | CGGACGGAGGGAGTATTTCT |
| Hv_aCENH3_EX7_R | GGACATGCCCAAAGAAAGTG |
| Hv_bCENH3_EX1+2_F | GCCAGCGAGTACTCCTACAAG |
| Hv_bCENH3_EX1+2_R | TTGAGTTACCAGCCACCACTC |
| Hv_bCENH3_EX3_F | GTCATGCACTGTGTCTTGCA |
| Hv_bCENH3_EX3_R | TGCTAAGATCGGATAACTGTGG |
| Hv_bCENH3_EX4_F | TGCTCCTGAACAAACTGAACC |
| Hv_bCENH3_EX4_R | GTGGCCGTCAGTACAATCG |
| GAPDH-F | CAATGATAGCTGCACCACCAACTG |
| GAPDH-R | CTAGCTGCCCTTCCACCTCTCCA |
| Hvα-F | AGTCGGTCAATTTTCTCATCCC |
| Hvα-R | CTCTGTAGCCTCTTGAACTGC |
| Hvß-F | GCCATTGTCGAACAAGAAGG |
| Hvß-R | TAACACGGTGCGAATGAATG |
| A.tha-CENH3-F | ATGGCGAGAACCAAGCATCG |
| A.ar-CENH3CDNA-R | TCACCATGGTCTGCCTTTTC |
| CH3A+L130_F_for | phos-GACAGCTGAAGCATTTGTTGCTCTTC |
| CENH3L130_I_for | phos-GACAGCTGAAGCTATTGTTGCTCTTC |
| CENH3L130_F+I_rev | phos-CAACGATTGATTTGGGGAGGG |
| cenh3-1_mut_for | GGTGCGATTTCTCCAGCAGTAAAAATC |
| cenh3-1_mut_rev | CTGAGAAGATGAAGCACCGGCGATAT |
| cenh3-1_mut2429r | AACTTTTGCCATCCTCGTTTCTGTT |
| BvCENH3-cds1 | GGATCCATGAGAGTTAAACACACTGC |
| BvCENH3-cds2 | GGATCCTGTTCAGTTACCATCCCCTC |
| BvCENH3_mut_Fw | ATGGATCCATGAGAGTTAAACACACTGC |
| BvCENH3_L->F_Rv | CTCTGCAGCCTCTTGAAGGGCCATAAAAGC |
| BvCENH3_L->I_Rv | CTCTGCAGCCTCTTGAAGGGCCATAATAGC |
RNA extraction, PCR, and quantitative real-time RT-PCR.
Total RNA was isolated from roots and leaves using the TRIzol method (16) and from anthers (microscopically staged between meiosis and development of mature pollen), carpel, endosperm, and embryo by a Picopure RNA isolation kit (Arcturus) according to the manufacturer’s instructions. Transcript levels of each gene were normalized to GAPDH by the following formula: R = 2^(-(CtGOI–CtH))*100 (17), where R = relative changes, GOI = the gene of interest, and H = housekeeping (GAPDH). The specificity and efficiency of both primers were determined by quantitative RT-PCR (qRT-PCR) using a dilution series of plasmids of cloned full-length barley αCENH3 and βCENH3 genes. A similar Ct value (the PCR cycle at which the fluorescent signal of the reporter dye exceeds background level) for equal amount of plasmid indicates that both primers can amplify specific transcripts with the same efficiency.
Indirect immunostaining.
Indirect immunostaining of nuclei and chromosomes was carried out as described in ref. 6. CENH3 of barley was detected with guinea pig anti-αCENH3– and rabbit anti-βCENH3–specific antibodies. A rabbit HTR12-specific antibody (ab72001; Abcam) was used for the detection of A. thaliana CENH3 (AtCENH3).
A. thaliana.
Cloning and generation of CENH3 transgenes.
To generate CENH3 genomic fragments carrying mutations, resulting in phenylalanine 130 (F130) and isoleucine 130 (I130) instead of wild-type leucine 130 (L130), a genomic CENH3 fragment in the pCAMBIA1300 vector used to complement cenh3-1/cenh3-1 (the cenh3-null mutant) (2, 18) was subcloned via the unique HindIII and BamHI sites into pBlueScript II KS (Strategene; www.genomics.agilent.com/). Mutations of CENH3 (L130I or L130F) were generated in pBlueScript II KS using a Phusion Site-Directed Mutagenesis Kit (Finnzymes; diagnostics.finnzymes.fi/reagents_index.html) according to the manufacturer’s instructions with minor changes as described in ref. 19. The following 5′-phosphorylated primers were used for mutagenesis: CH3A+L130_F_forward, CENH3L130_I_forward, and CENH3L130_F+I_reverse (Table S2). Mutated CENH3 genomic fragments were subcloned via the unique HindIII and BamHI sites into the initial pCAMBIA1300 vector (18) containing a hygromycin resistance marker. All constructs were verified by sequencing. To generate p35S::eYFP-CENH3 fusion constructs containing mutations within the CENH3 coding DNA sequence, resulting in L130I or L130F, a plasmid (p35S-BM; www.dna-cloning.com) containing a p35S::eYFP-CENH3 expression cassette (20) was used as template for the Phusion Site-Directed Mutagenesis Kit (Finnzymes; diagnostics.finnzymes.fi/reagents_index.html). The primers (Table S2) and strategies used to introduce the desired mutations were the same as above. The resulting expression cassettes [35Spro, eYFP-(mutated)CENH3, and NOS terminator] were subcloned via unique SfiI restriction sites into the pLH7000 vector containing a phosphinotricine resistance marker (www.dna-cloning.com) and were verified by sequencing.
Plant transformation, culture conditions, and cross-pollination.
A. thaliana wild-type and cenh3-1/CENH3 heterozygotes plants (both accession Columbia-0) were transformed by the floral dip method (21). Transgenic progenies were selected on MS (22) solid medium containing the corresponding antibiotic. Plants were germinated on Petri dishes under long-day conditions (20 °C, 16 h light/18 °C, 8 h dark), grown for 4 wk under short-day conditions (20 °C 8 h light/18 °C 16 h dark), and then shifted to long-day conditions again. For crossing, closed buds of the A. thaliana cenh3 mutant were emasculated by removing the immature anthers. The stigmas of emasculated buds were pollinated using the yellowish pollen from mature anthers of freshly opened, wild-type A. thaliana flowers.
DNA extraction and genotyping of A. thaliana.
Genomic DNA preparations and PCR-based genotyping were performed using standard methods. DNA was extracted according to ref. 23. Plants were genotyped for cenh3-1 in a derived cleaved amplified polymorphic sequence (dCAPS) genotyping reaction. The dCAPS primers cenh3-1_mut_forward and cenh3-1_mut_reverse were used to amplify CENH3. Amplicons were digested with EcoRV and resolved on a gel. The cenh3-1 mutant allele is not fragmented (215 bp), whereas the wild-type CENH3 allele is cut (191 and 24 bp). (For primers see Table S2.) To genotype the endogenous CENH3 locus for cenh3-1 in the offspring of cenh3-1/CENH3 plants transformed with the CENH3 genomic locus (untagged CENH3 transgene with L130, L130I, or L130F), an initial PCR was performed with one primer outside the transgene CENH3 locus, allowing specific amplification of the endogenous but not the transgenic CENH3 locus. Primers used were cenh3-1_mut_for and cenh3-1_mut2429r. Amplicons were purified and used as template for a second dCAPS PCR genotyping reaction as described above for cenh3-1 plants. The presence of the transgene was verified by sequencing of the RT-PCR product using the primer set of A.tha-CENH3-F and A.ar-CENH3CDNA-R (Table S2).
Western blot analysis.
Nuclear proteins were isolated (24) and separated in 10% polyacrylamide gels according to ref. 25. Samples were electrotransferred onto Immobilon PVDF membranes (Millipore). Membranes were simultaneously incubated for 12 h at 4 °C in PBS and 5% low-fat milk with histone H3 mouse monoclonal (1:5,000; Sino Biological Inc., catalog no.100005-MM01-50) and A. thaliana CENH3 (HTR12) rabbit polyclonal (1:5,000; ab72001; Abcam) primary antibodies. Anti-mouse IRDye 680RD (1:5,000; LI-COR) and anti-rabbit IRDye 800CW (1:5,000; LI-COR) were used as secondary antibodies. Signals were detected by the Odyssey CLx Imager (LI-COR). Histone H3 signals were used to equalize the amounts of loaded proteins.
Flow cytometric analysis of A. thaliana plants.
For flow cytometric ploidy analyses of plants, equal amounts of leaf material from 5–10 individuals were chopped together with a sharp razor blade in nuclei isolation buffer (26) supplemented with DNase-free RNase (50 µg/mL) and propidium iodide (50 µg/mL). The nuclei suspensions were filtered through a 35-µm cell-strainer cap into 5-mL polystyrene tubes (BD Biosciences) and measured on a FACStarPLUS cell sorter (BD Biosciences) equipped with an argon ion laser INNOVA 90C (Coherent). Approximately 10,000 nuclei were measured and analyzed using the software CELL Quest ver. 3.3 (BD Biosciences). The resulting histograms were compared with a reference histogram of a diploid wild-type plant. If an additional peak indicated haploidy, single plants were measured again to identify the haploid individuals.
Barley test crosses.
The barley TILLING line 4528 was first made homozygous for the leucine-to-phenylalanine substitution at amino acid 92 of βCENH3 via generational segregation. To reduce the mutation background, it then was crossed with wild-type barley cv. Golden Promise. As expected, all F1 hybrids carried the ßcenh3 mutation in a heterozygous state. Upon selfing of F1 plants and sequencing βCENH3 of F2 individuals, 14 selected homozygous βcenh3 segregants derived from seven independent F1 hybrids were used for reciprocal crosses with wild-type barley. To this end, the spikes used as maternal material were emasculated and bagged to prevent any unintended pollination. Mutant plants were used as the maternal parent in 36 crosses and as pollinator in 22 crosses. As a positive control for uniparental genome elimination, emasculated spikes of Golden Promise were pollinated by H. bulbosum (accessions HbPB1 and HbFBB of the Leibniz Institute of Plant Genetics and Crop Plant Research (IPK) GenBank). Cross-pollinated plants were transferred to the glasshouse with day/night temperatures of 20/18 °C, respectively (27). To stimulate caryopsis and embryo development, despite potential chromosome loss, 100 mg/L 2,4-D or dicamba was applied by injection into the uppermost internode of the spike or by dropping into the individual florets 1–4 d after pollination (28). Embryos were excised aseptically from the caryopses 18–21 d post pollination and placed with the scutellum facing down on B5 medium (29) to regenerate plantlets as described (30). Genome size measurements of plantlets established in soil were conducted using a Ploidy Analyzer I (Partec).
B. vulgaris.
Plant transformation and culture conditions.
Stable transformation of B. vulgaris callus was performed as described (31) by selection using kanamycin. After ∼2 mo [24 °C, 16 h light (55 µmol⋅m−2⋅s−1)/8 h dark], callus cells were microscopically analyzed.
Cloning and generation of CENH3 transgenes.
To generate the 35S::RFP-CENH3 fusion construct, CENH3 was amplified from sugar beet cDNA (BvCENH3-cds1 and BvCENH3-cds2) and cloned into a vector containing a 35Spro, RFP, and 35S-terminator expression cassette. For constructs containing mutations within the CENH3 coding sequence, resulting in F106 and I106 instead of L106, the above-mentioned plasmid containing the 35S::RFP-CENH3 expression cassette was used as template for primer-based mutagenesis. The PstI site close to the position of the desired mutation was used to split CENH3 into two parts. The desired mutations were integrated via mutations in the primers (BvCENH3_mut_Fw, BvCENH3_L->F_Rv, BvCENH3_L->I_Rv; Table S2). The resulting expression cassettes [35Spro, RFP-(mutated)CENH3 and 35S-terminator] were verified by sequencing. For further details see SI Materials and Methods.
SI Materials and Methods
RNA Extraction, PCR, and Real-Time qRT-PCR.
The absence of genomic DNA contamination was confirmed by PCR using GAPDH primers (Table S2). Ten microliters of PCR mixture contained 1 μL of cDNA template, 5 μL of 2× Power SYBR Green PCR Master Mix (Applied Biosystems), and 0.33 mM of the forward and reverse primers for each gene (Table S2). Reactions were run in an Applied Biosystems 7900HT Fast Real-Time PCR System. The PCR was performed using the following conditions: 95 °C for 10 min, followed by 40 cycles at 95 °C for 15 s, at the annealing temperature of 60 °C for 60 s. Three technical replicates were performed for each cDNA sample. Data were analyzed with SDS software v. 2.2.2.
Indirect Immunostaining.
Epifluorescence imaging was performed using an Olympus BX61 microscope and an ORCA-ER CCD camera (Hamamatsu). To analyze the structures of immunosignals and chromatin at an optical resolution of ∼120 nm (superresolution), structured illumination microscopy (SIM) was applied using a C-Apo 63×/1.2W Korr objective of an Elyra PS.1 microscope system and the ZEN software (Zeiss). Images were captured separately for each fluorochrome using the 561, 488, and 405-nm laser lines for excitation and appropriate emission filters.
Genotyping of A. thaliana.
To determine the copy number of transgenes, EcoR1-digested genomic DNA of A. thaliana was gel separated and transferred to a Hybond XL membrane (GE Healthcare Life Sciences). Southern hybridization was performed with the ɣ32P-labeled hptII (hygromycin resistance) gene. After hybridization, membranes were exposed overnight to a Fuji phosphoimager screen. Signals were detected using a Fuji phospho imager (FLA-3000).
Analysis of CENH3 Localization in B. vulgaris.
To analyze the localization of CENH3 and the mutated CENH3 callus, material was analyzed using a C-Apo 63×/1.2W Korr objective of an Axio Imager M2 microscope system and ZEN software (Zeiss).
Acknowledgments
We thank the late Simon Chan (University of California, Davis) for sharing the A. thaliana cloned CENH3 genomic fragment and A. thaliana cenh3-null mutant plants; Inna Lermontova [Leibniz Institute of Plant Genetics and Crop Plant Research (IPK)] for providing the p35S::eYFP-CENH3 fusion construct; Twan Rutten (IPK) for scanning electron microscopy; Ingo Schubert and Florian Mette (IPK) for stimulating discussions; and Katrin Kumke, Karla Meier, Heike Büchner, Petra Hoffmeister, Jacqueline Pohl, and Marzena Kurowska for excellent technical assistance. This work was supported by the German Federal Ministry of Education and Research Plant 2030 Project HAPLOIDS FKZ 0315965, FKZ0313123C, and FKZ0315052B and the German Research Foundation (DFG) Collaborative Research Center 648.
Footnotes
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1504333112/-/DCSupplemental.
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